ES2829568T3 - Biomedical patches with aligned fibers - Google Patents

Abstract

A method of producing a fiber structure, the method comprising: electrically charging one or more first electrodes defining an area with a first polarity; electrically charging a second electrode located within the area defined with the first polarity; electrically charging a row with a second polarity opposite the first polarity, wherein the row is aligned with the second electrode; and dispensing a polymer from the spinneret to create a fiber structure extending from the second electrode to the first electrode (s).

Description

DESCRIPTION

Biomedical patches with aligned fibers

Background

Many surgical procedures result in the perforation or removal of biological tissue, such as the water-impermeable fibrous membrane that surrounds the brain, known as the dura. In some cases, for example minimally invasive neurosurgical procedures, relatively few small holes are created in the dura, while in others, for example surgical resection of advanced tumors, large sections of the dura can be removed. dura mater. In all of these cases, the tissue barrier surrounding the brain must be repaired to prevent damage to cortical tissues and leakage of cerebrospinal fluid. To facilitate this repair, neurosurgeons use sheets of polymeric materials or processed tissue, which act like native dura, and are known as dural substitutes.

At least some known dural substitutes used in neurosurgical clinics are composed of an acellular collagen matrix obtained from isolated bovine or porcine tissues. Although generally accepted in this field, such xenogeneic dural substitutes can increase the incidence of adhesions and contractures, transmit various zoonotic diseases to patients, and generally adversely affect the outcome of patients after surgery. Additionally, processed collagen grafts are extremely expensive, costing patients and insurance companies thousands of dollars per procedure.

Furthermore, although cell microarrays can be useful in biomedical research and tissue engineering, at least some known techniques for the production of such cell microarrays can be expensive and require a long period of time, and can also require the use of specialized instruments and sophisticated.

A method and apparatus for producing nanofibers are disclosed in Japanese Patent No. 2008223186 A, by which the orientation of accumulated nanofibers in a collector can be improved.

A method and apparatus for the production of patterned and textured nonwoven fabrics are disclosed in US Patent No. 3,280,229 A, wherein electrically spun filaments are aggregated into bundles or groups that define a controlled pattern of aggregates of high density of filaments.

A method and apparatus for forming electrospun fibers which are deposited on a collecting surface to thereby form a patterned polymer structure are disclosed in US Patent No. 2005/104258 A1.

An apparatus and method for electrospinning under a first electrical polarity first fibers of a first substance, electrospinning under a second electrical polarity fibers of a second substance and joining the first and second fibers are disclosed in US Patent No. 2006/264140 A1. to form a fiber mat.

Summary

In accordance with one aspect of the invention, a method for producing a fiber structure is disclosed, and this method comprises electrically charging one or more first electrodes defining an area with a first polarity, electrically charging a second electrode located within the defined area with the first polarity, electrically charging a die (spinneret) with a second polarity opposite the first polarity, wherein the die is aligned with the second electrode, and dispensing a polymer from the die to create a structure of fibers extends from the second electrode to the one or more first electrodes.

In accordance with another aspect of the invention, an apparatus is disclosed for producing a structure that includes a plurality of fibers, this apparatus comprising a plurality of first electrodes at least partially circumscribing an area, a second electrode located within the area, and electrically coupled to the first electrodes and a power source electrically coupled to the first electrodes and the second electrode, wherein the power source is configured to electrically charge the first electrodes and the second electrode with a first polarity, wherein when a row aligned with the second electrode and charged with a second polarity opposite the first polarity dispenses a polymer, the dispensed polymer forms a plurality of fibers extending from the second electrode to the first electrodes.

One or more disclosures described herein provide structures that have a plurality of aligned fibers (eg, radially aligned and / or polygonally aligned). When such a structure is used as a biomedical patch, the alignment of the fibers, as described herein, can provide directional cues that influence cell spread. For example, the structures provided can promote new cell growth along the fibers, so that cell propagation in one or more desired directions can be obtained.

One or more disclosed structures provided can be created using an apparatus that includes one or more first electrodes that define an area and / or partially circumscribe an area. For example, a single first electrode can enclose the area, or a plurality of one or more first electrodes can be positioned on at least a portion of the perimeter of the area. A second electrode is placed within the area. In exemplary disclosures, when the electrodes are electrically charged with a first polarity, and a spinneret dispensing a polymer (for example, toward the second electrode) is electrically charged with a second polarity opposite the first polarity, the dispensed polymer forms a plurality of fibers extending from the second electrode to the first electrodes. Furthermore, electrodes with rounded (eg, convex) surfaces can be configured in an array, and a fibrous structure created using such electrodes can include an array of wells at positions corresponding to the positions of the electrodes.

The scope of protection is defined by the claims. Any subject matter outside the scope of the claims is provided for comparative purposes only.

Brief description of the drawings

The embodiments described herein can be better understood by referring to the following description, accompanied by the accompanying drawings.

Figure 1 is a diagram illustrating a perspective view of an example of an electrospinning system for producing a radially aligned fiber structure.

Figure 2 is a diagram illustrating an electric field generated by the electrospinning system shown in Figure 1.

Figure 3 is a diagram of an electrode removed from the electrospinning system shown in Figure 1 and having a plurality of fibers deposited thereon that form a biomedical patch.

Figure 4 is a photograph of a biomedical patch that includes a plurality of radially aligned electrospun fibers deposited on a peripheral electrode.

Figure 5 is a scanning electron microscope (SEM) image of the biomedical patch shown in Figure 4, further illustrating that the fibers of the biomedical patch are radially aligned.

Figure 6 is an illustration of a row of solid fibers.

Figure 7 is an illustration of a hollow fiber spinneret.

Figure 8 is an illustration of a biomedical patch layer with a plurality of randomly oriented fibers, a biomedical patch layer with a plurality of radially aligned fibers, and a multilayer biomedical patch that includes multiple arrays of fibers.

Figure 9 is a diagram of a collector with a center electrode, an inner peripheral electrode defining an inner closed area, and an outer peripheral electrode defining an outer closed area.

Figure 10 is a diagram of a concentric biomedical patch that can be produced using the manifold shown in Figure 9 in conjunction with the electrospinning system shown in Figure 1.

Figure 11 is a flow chart of an exemplary method for producing a radially aligned fiber structure using a peripheral electrode defining a closed area and a center electrode located approximately in a center of the closed area.

Figure 12 is a flow chart of an exemplary method for repairing a defect, damage, or void in biological tissue.

Figure 13 is a schematic illustration of cellular infiltration of a biomedical patch from intact dural tissue adjacent to the edge of a biomedical patch.

Figures 14A-14D are fluorescence micrographs comparing cell migration when dura tissues were grown on scaffolds of radially aligned nanofibers and randomly oriented nanofibers for 4 days.

Figure 15 is a schematic diagram of a cell culture system custom designed to model the wound healing response of defects or voids in biological tissue.

Figures 16A-16D are fluorescence micrographs showing the morphology and distribution of cells in scaffolds of radially aligned nanofibers and randomly oriented nanofibers with and without fibronectin coating after incubation for 1 day.

Figures 17A-17D are fluorescence micrographs showing the migration of dura fibroblasts seeded on radially aligned nanofiber fibronectin-coated scaffolds for 1 day, 3 days, and 7 days.

Figure 18 is an illustration of a method used to determine the area of the remaining acellular region of the nanofiber scaffolds within the simulated tissue defect.

Figure 19 is a graph illustrating the remaining acellular area in the nanofiber scaffold within the simulated tissue defect as a function of incubation time.

Figures 20A-20D are fluorescence micrographs showing membrane dye-labeled live dural fibroblasts on radially aligned nanofiber scaffolds coated with fibronectin after a 1-day culture, a 3-day culture, a fibronectin culture. 7 days and a 10-day culture.

Figures 21A-21D are fluorescence micrographs demonstrating the organization of cells and adherent extracellular matrix into scaffolds of radially aligned fibers and randomly oriented fibers obtained by immunostaining for type I collagen (green) and cell nuclei (blue).

Figure 22 is a graph illustrating the thickness of the regenerated dura at the center of repaired dural defects over time.

Figure 23 is a graph illustrating the content of regenerative collagenous tissue over time. Figure 24 is a diagram illustrating a perspective view of an exemplary electrospinning system for producing a polygon-aligned fiber structure using an electrode array.

Figure 25 is a diagram illustrating an elevational view of an example of a modular electrospinning collector. Figure 26 is a diagram illustrating an electric field generated by an electrospinning system such as the electrospinning system shown in Figure 24.

Figures 27A-27F are microscopy images of a membrane produced using a collector with an electrode array, such as the collector shown in Figure 24.

Figures 28A-28D are microscopy images illustrating cell growth on a membrane as shown in Figures 27A-27F.

Figures 29A and 29B are microscopy images illustrating neurite propagation in a membrane as shown in Figures 27A-27F.

Figures 30A and 30B are overlays of a light microscopy image and a fluorescent microscopy image illustrating the formation of neural networks from embryoid bodies in a membrane as shown in Figures 27A-27F.

Figures 31A-31D are scanning electron microscopy images illustrating membranes produced using a variety of electrode arrays.

Figure 32 is a diagram of a collector with peripheral electrodes partially circumscribing an area. Detailed description

The embodiments disclosed herein facilitate the repair of biological tissue with the use of a biomedical patch that includes a plurality of fibers. These fibers can have a very small cross-sectional diameter (eg, 1-1000 nanometers) and consequently can be called nanofibers. Although biomedical patches are described herein with reference to the dura and its use as a dural substitute, the described embodiments can be applied to any biological tissue. Also, although they are described as biomedical patches, the structures with aligned fibers can be used for other purposes. Accordingly, the described embodiments are not limited to biomedical patches.

During their operation, the biomedical patches disclosed herein facilitate cell growth and may be referred to as "membranes,""scaffolds,""matrices," or "substrates." Such biomedical patches also facilitate cell migration from a perimeter of the patch to a center of the biomedical patch. The patches Fiber-aligned biomedicals, as described herein, can promote significantly faster healing and / or regeneration of tissue, such as the dura, than substitutes, which lack nanoscopic organization and directional cues.

The dura is a membranous connective tissue located in the outermost layer of the three layers of the meninges that surround the brain and spinal cord, which covers and supports the dural sinuses and carries blood from the brain to the heart. Dural substitutes are often required after a neurosurgical procedure to repair, expand, or replace the cut, damaged, or resected dura.

Although many efforts have been made in this regard, the challenge of developing an appropriate dural replacement has met with only limited success. Autografts (eg, fascia lata, temporal fascia, and pericranium) are preferable because they do not cause severe inflammatory or immune reactions. Potential drawbacks of autografts include difficulty in achieving a waterproof closure, scar tissue formation, insufficiently accessible graft materials to close large dura defects, increased risk of infection, site morbidity donor and the need for an additional operating site. Allografts and xenografts are often associated with adverse reactions such as graft dissolution, encapsulation, foreign body rejection, scarring, adhesion formation, and toxicity side effects resulting from immunosuppressive regimens. Freeze-dried human dura as a dural substitute has also been reported to be a source of communicable diseases specifically related to prions, such as Creutzfeldt-Jakob disease.

In terms of materials, nonabsorbable synthetic polymers, such as silicone and expanded polytetrafluoroethylene (ePTFE), often cause serious complications that may include induction of granulation tissue formation due to chronic stimulation of the response to foreign body. Natural absorbable polymers, such as collagen, fibrin, and cellulose, can pose a risk of infection and disease transmission. As a result, synthetic polymers such as poly (3-hydroxybutyrate-co-3-hydroxyvalerate) (PHBV), polylactic acid (PLA), polyglycolic acid (PGA), ternary copolymers PLA-PCL-PGA, and hydrogels of Hydroxyethylmethacrylate has recently sparked interest as biodegradable implant materials for dura repair. The methods and systems described herein can be practiced with these materials and / or with any biomedical polymer.

In order to facilitate the successful regeneration and / or repair of the dura after a surgical operation, a biomedical patch or synthetic dural substitute should promote: (i) the adhesion of dural fibroblasts (the type of primary cell present in the dura mater) to the surface of the biomedical patch; (ii) migration of dural fibroblasts from the periphery of the biomedical patch towards the center; and (iii) a minimal immune response. To date, only synthetic dural substitutes in the form of sheets, films, mesh, glues and hydrogels have been tested. Due to the isotropic surface properties, such substitutes are not very suitable for cell attachment and inward directed migration.

This problem can potentially be solved by manufacturing the polymers as nanoscale fibers with a specific order and organization. For example, the cell migration rate can be very low on flat isotropic surfaces, whereas cells can migrate a very long distance in a highly correlated manner with a constant speed in a fibrous and uniaxially aligned scaffold.

Electrospinning is a capacitive technique that can produce nanoscale fibers from a large number of polymers. Electrospun nanofibers are typically collected as a randomly oriented nonwoven mat. Uniaxially aligned nanofiber arrays can also be obtained under certain conditions, specifically when using an air chamber manifold or high speed rotating mandrel. However, uniaxially aligned nanofiber scaffolds promote cell migration only along a specific direction and are therefore not ideal as dural substitutes.

In order to promote cell migration from the surrounding tissue to the center of a dural defect and reduce the healing and regeneration time of the dura, a patterned surface with aligned nanoscale features (for example, aligned radially and / or in one or more polygons) would be very advantageous as an artificial dural substitute. More specifically, scaffolds constructed of aligned nanofibers could meet this demand by guiding and enhancing cell migration from the edge of a dural defect to the center.

There are a large number of polymers available for use in electrospinning. In some embodiments described herein, nanofibers for dural substitutes such as electrospun polymer are produced from poly (scaprolactone) (PCL), an FDA-approved semi-crystalline polyester that can degrade by hydrolysis of their ester bonds under physiological conditions with non-toxic degradation products. This polymer has been widely used and studied in the human body as a material for the manufacture of drug delivery carriers, sutures or adhesion barriers. As described herein, electrospun PCL nanofibers can be aligned to generate scaffolds that are useful as dural substitutes.

The embodiments disclosed herein facilitate the production of a new type of artificial tissue substitute that includes a polymeric nanofiber material, which is formed through a new electrospinning method. This polymeric material includes nonwoven nanofibers (eg, fibers having a diameter of 1-1000 nanometers) that are aligned within a sheet of material.

In exemplary embodiments, a material with aligned nanofibers is formed through a novel electrospinning method that uses a collector that includes one or more first electrodes, or "peripheral" electrodes, that define an area and / or at least partially circumscribe said area. area, and a second "internal" electrode located within the area. When the electrodes are electrically charged with a first polarity, and a spinneret that dispenses a polymer (for example, towards the inner electrode) is electrically charged with a second polarity opposite the first polarity, the dispensed polymer forms a plurality of fibers that are extend from the inner electrode to the peripheral electrode (s). The electrodes may include a rounded (eg, convex) surface so that a depression or "well" is formed on the electrode-facing side of a fiber structure. Alternatively, the electrodes may include a concave surface, such that a well is formed on the side of the frame opposite the electrodes.

In some embodiments, the collector includes a single inner electrode and a single peripheral electrode. In other embodiments, the collector includes a plurality of peripheral electrodes and the dispensed polymer may form fibers that extend between said peripheral electrodes, in addition to fibers that extend from the inner electrode to one or more of the peripheral electrodes.

Furthermore, in some embodiments, there are multiple areas defined and / or partially circumscribed by peripheral electrodes. For example, an inner peripheral electrode can define an inner closed area that surrounds the inner electrode, and an outer peripheral electrode can define an outer closed area that surrounds the inner peripheral electrode. In other embodiments, the electrodes are configured in a matrix, such as a grid and / or other polygonal pattern (eg, a hexagonal pattern), and multiple partially overlapping areas can be defined by said electrodes. For example, an internal electrode of one area can function as a peripheral electrode of another area. In such embodiments, the dispensed polymer can form fibers that extend between the collector electrodes, such that the fibers define the sides of a plurality of polygons, with the electrodes located at the vertices of the polygons.

Unlike known nanofiber structures, the aligned nanofiber materials disclosed herein are capable of presenting nanoscale topographic indications to local cells that enhance and direct cell migration (e.g., throughout the sheet of material or into the center of the media sheet). As a result, aligned nanofiber materials can cause faster cell population and migration than randomly oriented materials, such as reference processed collagen matrices. The materials described herein can be particularly useful as a substrate for various types of biomedical patches or grafts designed to induce protection, closure, healing, wound repair, and / or tissue regeneration.

An aligned nanofiber scaffold, as described herein, possesses significant potential as an artificial dural substitute, as it is capable of stimulating robust cell migration from the adjacent intact dura and promoting the rapid cell population of the required nanofiber matrix. to cause dural repair. Furthermore, said nanofiber materials offer the advantage of being economical to produce and of being fully compliant and resorbable. Dural nanofiber substitutes can also reduce the risk of contractures and completely eliminate the risk of transmitted zoonotic diseases when applied intraoperatively, generally improving patient outcomes after surgery.

Inner electrode and peripheral electrode (s)

Figure 1 is a diagram illustrating a perspective view of an exemplary electrospinning system 100 for producing a radially aligned fiber structure. System 100 includes a collector 105 with a first electrode 110, which may be referred to as a peripheral electrode, and a second electrode 115, which may be referred to as an inner or center electrode. The system 100 also includes a row 120. The peripheral electrode 110 defines a closed area 125 and the center electrode 115 is located approximately in a center of the closed area 125.

System 100 is configured to create an electrical potential between collector 105 and row 120. In one embodiment, peripheral electrode 110 and center electrode 115 are configured to be electrically charged with a first amplitude and / or polarity. For example, peripheral electrode 110 and center electrode 115 may be electrically coupled to power source 130 through conductor 135. Power source 130 is configured to charge peripheral electrode 110 and center electrode 115 with the former. amplitude and / or polarity across lead 135.

In the embodiment illustrated in Figure 1, peripheral electrode 110 is a ring defining a closed area 125 that is circular. For example, the circular closed area 125 may have a diameter of between 1 cm and 20 cm. In other embodiments, the peripheral electrode 110 may have any shape suitable for use with the described methods. at the moment. For example, peripheral electrode 110 may define a closed area 125 elliptical, ovular, rectangular, square, triangular, and / or otherwise rectilinear or curvilinear. In some embodiments, peripheral electrode 110 defines a closed area 125 of between 5 cm2 and 100 cm2. Peripheral electrode 110 may have a height 112 of between 0.5 cm and 2.0 cm. The center electrode 115 may include a metallic needle and / or any other structure that ends in a point or set of points.

In one embodiment, closed area 125 defines a horizontal plane 127. Row 120 is aligned with center electrode 115 and is vertically offset from horizontal plane 127 by a variable distance. For example, row 120 may be offset vertically with respect to horizontal plane 127 by a distance of between 1 cm and 100 cm.

Spinneret 120 is configured to dispense polymer 140 while electrically charged with a second amplitude and / or polarity opposite the first polarity. As shown in Figure 1, string 120 is electrically coupled to power source 130 via lead 145. Power source 130 is configured to charge string 120 with the second amplitude and / or polarity through lead 145 In some embodiments, the power supply 130 provides a direct current (DC) voltage (eg, between 10 kilovolts and 17 kilovolts). In one embodiment, conductor 145 is positively charged, and conductor 135 is negatively charged or grounded. In some embodiments, power supply 130 is configured to allow adjustment of a current, a voltage, and / or a power.

In one embodiment, the spinneret 120 is coupled to a syringe 150 containing a polymer 140 in the form of a liquid solution. Syringe 150 can be operated manually or by a syringe pump 155. In one exemplary embodiment, spinneret 120 is a metal needle having an aperture between 100 microns and 2 millimeters in diameter.

As syringe 150 exerts pressure on polymer 140, spinneret 120 dispenses polymer 140 in the form of flow 160. Flow 160 has a diameter approximately equal to the diameter of the opening of spinneret 120. Flow 160 descends toward the collector 105. For example, flow 160 may fall downward under the influence of gravity and / or may be drawn downward by a charged conductive surface 162 located below collector 105. For example, conductive surface 162 may be electrically coupled to conductor 135 and charged with the same amplitude and / or polarity as peripheral electrode 110 and center electrode 115. As flow 160 descends, polymer 140 forms one or more solid polymeric fibers 165.

In some embodiments, a mask 164, composed of a conductive or non-conductive material, is applied to collector 105 to manipulate fiber deposition 165. For example, mask 164 can be positioned between row 120 and collector 105 in such a way that no fibers 165 are deposited in collector 105 below mask 164. In addition, mask 164 can be used as a time-varying mask by adjusting its position while spinneret 120 dispenses polymer 140, facilitating spatial variation in density. of fibers in collector 105. Although mask 164 is shown as circular, mask 164 may be of any shape (eg, rectangular or semi-circular) and of a size suitable for use with system 100. Alternatively or additionally, the Deposition of fibers 165 in collector 105 can be manipulated by adjusting the position of collector 105 relative to row 120 or by spatially varying the electrical potential applied between row 120 and / or or the electrodes that make up collector 105. For example, the location of one side of collector 105 directly below row 120 may cause more fibers 165 to be deposited on that side than are deposited on the opposite side of collector 105.

Figure 2 is a diagram 200 illustrating an electric field generated by system 100. Diagram 200 shows a two-dimensional cross-sectional view of electric field force vectors between row 120 and peripheral electrode 110 and center electrode. 115 from manifold 105 (shown in Figure 1). Unlike known electrospinning systems, the electric field vectors (flux lines) in the vicinity of the collector are divided into two populations, those pointing towards the peripheral electrode 110 and those pointing towards the central electrode 115.

If the effect of the loads on the polymeric fibers is not found, the potential field v V = zJL can be calculated

using the Poisson equation, e , where V is the electric potential, £ is the electric permittivity of air, and P is the space charge density. The electric field, E, can be calculated by taking the negative gradient of the electric potential field, E - —V V Here, the electric field was calculated to verify the alignment effect demonstrated by the deposited fibers, which was carried out using the software COMSOL3.3.

Figure 3 is a diagram of peripheral electrode 110 removed from electrospinning system 100 (shown in Figure 1) and having a plurality of fibers 165 deposited on top of it forming a biomedical patch 170. Fibers 165 extend radially between a center 175 corresponding to the position of the center electrode 115 (shown in Figure 1) and a perimeter 178 corresponding to the position of the peripheral electrode 110. For example, the perimeter 178 may be a circular perimeter around the center 175 that defines a diameter of between 1 cm and 6 cm.

Biomedical patch 170 is illustrated with a small number of fibers 165 in Figure 3 for clarity. In some embodiments, biomedical patch 170 includes thousands, tens of thousands, hundreds of thousands, or more of fibers 165, uniformly distributed throughout the closed area 125 (shown in Figure 1) of peripheral electrode 110. Even with millions of fibers 165, the biomedical patch 170 is flexible and / or malleable, which facilitates the application of the biomedical patch 170 on uneven surfaces of biological tissue, such as the surface of the dura mater.

The radial alignment of the fibers 165 demonstrates the shortest possible path between the perimeter 178 and the center 175. Consequently, the biomedical patch 170 also facilitates cell migration directly from the perimeter 178 to the center 175, allowing a reduction in the time required to the infiltration and population of cells in the applied biomedical patch, as well as for the regeneration of native tissue.

Fibers 165 have a diameter of 1-1000 nanometers. In one embodiment, the fibers have a diameter of about 220 nanometers (eg, 215 nm to 225 nm). The diameter of the fibers 165, the thickness of the biomedical patch 170, and / or the density of the fibers within the biomedical patch 170 can affect the durability (eg, tensile strength) of the biomedical patch 170. The biomedical patch can be produced. 170 with different mechanical properties by modifying the thickness and / or density of the fibers of the biomedical patch 170 by operating the electrospinning system 100 for relatively longer or shorter periods.

Figure 4 is a photograph 300 of a biomedical patch 305 that includes a plurality of radially aligned electrospun fibers deposited on a peripheral electrode 110. Figure 5 is a scanning electron microscope (SEM) 310 image of the biomedical patch 305, at which is further illustrated that the fibers of the biomedical patch 305 are radially aligned.

With reference to Figures 1 and 3, the fibers 165 can be solid or hollow. In some embodiments, the size and / or structure of the fibers 165 is determined by the design of the spinneret 120. Figure 6 is an illustration of a solid fiber spinneret 120A. The string of solid fibers 120A includes a conical body 180 that defines a center line 182. At a dispensing end 184, the conical body 180 includes a ring 186. The ring 186 defines a circular opening 190A through which it can be dispensed polymer 140. Fibers 165 produced with a row of solid fibers 120A have a solid composition.

Figure 7 is an illustration of a hollow fiber spinneret 120B. Like the solid fiber string 120A, the hollow fiber string 120B includes a conical body 180 with a ring 186 at a dispensing end 184. The hollow fiber string 120B also includes a central body 188B located within the ring 186. ring 186 and central body 188B define an annular opening 190B. Consequently, when polymer 140 is dispensed through hollow fiber row 120B, fibers 165 have a hollow composition, with an outer wall surrounding a cavity. The outer wall of a fiber 165 dispensed by the hollow fiber row 120B defines an outer diameter that corresponds to the inner diameter of ring 186 and an inner diameter that corresponds to the diameter of central body 188B. Accordingly, the outer diameter and the inner diameter of the hollow fibers 165 can be adjusted by adjusting the diameters of the ring 186 and the central body 188B.

The hollow fiber strand 120B facilitates the incorporation of a substance, such as a biological agent, growth factor, and / or drug (eg, a chemotherapeutic substance), into the biomedical patch 170. For example, the substance may be deposited within a cavity defined by hollow fibers 165 of biomedical patch 170. In one embodiment, polymer 140 is selected to create porous and / or semi-soluble fibers 165, and the substance is dispensed from the cavity through fibers 165. In another embodiment, polymer 140 is degradable and the substance is dispensed as fibers 165 degrade in vivo. For example, fibers 165 can be configured to degrade in 12 months, 6 months, or 3 months. The rate of degradation of polymer 140 can be manipulated by adjusting the proportion of the constituent polymers within polymer 140.

In another embodiment, solid fibers 165 deliver a substance. For example, a solid fiber 165 can be created from a polymer 140 that includes the substance in solution. As the solid fiber 165 degrades, the substance is released into the surrounding tissue.

As shown in Figures 6 and 7, the ring 186 is perpendicular to the center line 182. In an alternative embodiment, the ring 186 is oblique (for example, it is oriented at an acute or obtuse angle) with respect to the center line. 182. The outside diameter of the fibers 165 can be determined by the inside diameter of the ring 186.

Some embodiments facilitate the production of a biomedical patch that has radially aligned fibers and non-radially aligned fibers. For example, radially aligned fibers can be deposited in a first layer, and non-radially aligned fibers can be deposited in a second layer. Alternatively, the radially aligned and non-radially aligned fibers can be deposited in a single layer (eg, simultaneously, sequentially, and / or alternately). Referring to Figure 1, system 100 can be used to create randomly oriented fibers by charging or grounding conductive surface 162. Optionally, the Peripheral electrode 110 and center electrode 115 may be uncharged or ungrounded (eg, decoupled from conductor 135).

Figure 8 is an illustration of a biomedical patch layer 400 with a plurality of randomly oriented fibers 405 and a biomedical patch layer 410 with a plurality of radially aligned fibers 415. As shown in Figure 8, the layers (400 and 410) of biomedical patch can be combined (eg, overlapping) to produce a multilayer biomedical patch 420 with randomly oriented fibers 405 and radially aligned fibers 415, or any other combination of any number or type of fiber layers. The combination of non-radially aligned fibers 405 and radially aligned fibers 415 facilitates the delivery of a biomedical patch that promotes cell migration to a center of the biomedical patch, while also exhibiting potentially greater durability (e.g., tensile strength) than that of a biomedical patch having only radially aligned fibers 415. The combination of non-radially aligned fibers 405 and radially aligned fibers 415 can also allow spatial control of cell migration and infiltration along an axis perpendicular to the plane of the patch biomedical, thus facilitating the formation and organization of specific layers of cells and / or extracellular matrix proteins that resemble the layers of natural tissues.

In some embodiments, multiple layers 410 of biomedical patch can be combined with radially aligned fibers 415 to create a multi-layer biomedical patch. For example, referring to Figures 1 and 3, after depositing a first set of fibers in collector 105, one can wait for the first set of fibers 165 to fully solidify or cure and then deposit a second set of fibers 165 in collector 105. The second set of fibers 165 can be deposited directly on the first set of fibers 165 in collector 105. Alternatively, the first set of fibers 165 can be removed from collector 105 and the second set of fibers of fibers 165 can be deposited on conductive surface 162 and / or collector 105, and then removed and superimposed on first set of fibers 165. Such embodiments facilitate increased durability of the biomedical patch and additional spatial control of migration / cellular activity, even when only radially aligned fibers are used. In some embodiments, a hydrogel or polymeric scaffold may be placed between the biomedical patch layers 400 and / or the biomedical patch layers 410.

A multilayer biomedical patch can be useful for dural grafts and other tissue engineering applications. Sequential layers of fibers can be created with varying orders (for example, radially aligned or randomly oriented) and densities (for example, high or low fiber density), which can allow infiltration and population of specific cell types into selected layers of the artificial biomedical patch. For example, biomedical patches containing high fiber density generally prevent cell migration and infiltration, while biomedical patches containing low fiber density generally improve cell migration and infiltration.

In general, the ability to form multilayer fiber materials, as described herein, can be highly beneficial in the construction of biomedical patches designed to reproduce the natural multilayer structure of not only the dura, but also of other biological tissues such as skin, heart valve leaflets, the pericardium and / or any other biological tissue. Also, one or more layers of a biomedical patch can be made from biodegradable polymers, so that the resulting nanofiber materials are fully resorbed after implantation. Manipulation of the chemical composition of the polymers used to make these scaffolds can further allow specific control of the rate of degradation and / or reabsorption of a biomedical patch after implantation.

Some embodiments provide a biomedical patch that includes a plurality of nested (eg, concentric) areas. Figure 9 is a diagram of a collector 505 with a center electrode 115, a first or inner peripheral electrode 510 defining a first or inner closed area 515, and a second or outer peripheral electrode 520 defining a second or outer closed area 525 which is larger than the inner closed area 515. In some embodiments, the outer peripheral electrode 520 is oriented concentrically with the inner peripheral electrode 510. Although the inner peripheral electrode 510 and the outer peripheral electrode 520 are shown as defining closed circular areas 515, 525 In Figure 9, the inner peripheral electrode 510 and the outer peripheral electrode 520 may define closed areas 515, 525 in any way suitable for use with the methods described herein. Furthermore, the inner closed area 515 and the outer closed area 525 can be of different shapes and / or have different centers.

In operation with electrospinning system 100 (shown in Figure 1), central electrode 115 and inner peripheral collector 505 are charged with the first amplitude and / or polarity (opposite to the polarity with which row 120 is charged) , while spinneret 120 dispenses polymer 140 as flow 160. Flow 160 descends into manifold 505 and forms one or more fibers 530 extending from central electrode 115 to inner peripheral electrode 510.

The first polarity of the center electrode 115 is charged (for example, by decoupling the center electrode 115 from the lead 135), and the outer peripheral electrode 520 is charged with the first amplitude and / or polarity. Spinneret 120 dispenses polymer 140 as flow 160, which descends into collector 505 and forms one or more fibers 535 that extend from inner peripheral electrode 510 to outer peripheral electrode 520. Together, the fibers 530 and 535 form a concentric biomedical patch 550, as shown in Figure 10. In some embodiments, the charge is not removed from the central electrode 115 prior to depositing the fibers 535 between the inner peripheral electrode 510 and the outer peripheral electrode 520.

Figure 10 is a diagram of a concentric biomedical patch 550 that can be produced with the manifold 505 (shown in Figure 9). Fibers 530 define an inner area 555, shown as a circle extending from a center 560 to an inner perimeter 565. An outer area 570 includes fibers 535 that extend approximately from inner perimeter 565 (eg, approximately 100 pm to 2000 pm within the inner perimeter 565) to an outer perimeter 575. The fibers 535 are oriented radially or approximately (eg, 1, 3 or 5 degrees) radially with respect to the center 560.

As shown in Figure 10, the inner area 555 and the outer area 570 may overlap in an overlap area 580. In one embodiment, the overlap area 580 corresponds to a thickness of the inner peripheral ring 510 (shown in Figure 8 ). Similar to Figure 3, for clarity, Figure 10 shows concentric biomedical patch 550 with a small number of fibers 530 and 535. In some embodiments, inner area 555 and outer area 570 each include thousands , tens of thousands, hundreds of thousands or more fibers 530 and 535, respectively. Fibers 530 and fibers 535 can be coupled to each other in overlap area 580. For example, fibers 535 can be deposited before fibers 530 have fully solidified (or vice versa). In some embodiments, fibers 530 and fibers 535 are deposited in collector 505 (shown in Figure 9) simultaneously or alternately.

Embodiments such as those shown in Figures 9 and 10 facilitate the provision of a biomedical patch that has a relatively uniform fiber density throughout. In contrast, if fibers 530 were extended from the center 560 to the outer perimeter 575, the fiber density in the center 560 would be considerably higher than the fiber density in the outer perimeter 575. Low peripheral fiber density can compromise durability of a biomedical patch near an outer perimeter, especially in large diameters (for example, greater than 5 or 6 cm). Accordingly, such embodiments also facilitate the provision of a large diameter biomedical patch (eg, up to 10 or 12 cm), while maintaining the durability of the biomedical patch. In some embodiments, a non-radially aligned fiber layer is combined with the biomedical patch 550, as previously described with respect to Figure 8, which can further enhance the durability of the biomedical patch 550.

In some embodiments, the spatial density of the fibers within the inner area 555 is different from the spatial density of the fibers within the outer area 570. In one example, the fibers 530 are deposited between the center electrode 115 and the inner peripheral electrode 510 during a first period, and the fibers 535 are deposited between the inner peripheral electrode 510 and the outer peripheral electrode 520 during a second period.

Although collector 505 and concentric biomedical patch 550 are illustrated with circular inner and outer areas, any number and shape of peripheral electrodes can be used to create any number of distinct fiber areas within a biomedical patch.

Figure 11 is a flow chart of an exemplary method 600 for producing a radially aligned fiber structure using a peripheral electrode defining a closed area and a center electrode located approximately in a center of the closed area. Although one embodiment of method 600 is shown in Figure 11, it is envisioned that any of the illustrated operations may be omitted and that the operations may be performed in a different order than shown.

The method 600 includes electrically charging 605 the peripheral electrode and the center electrode with a first amplitude and / or polarity (eg, negatively charged or grounded). A row roughly aligned with the center electrode is electrically charged 610 with a second amplitude and / or polarity opposite the first amplitude and / or polarity (eg, with a positive charge).

A polymer (eg, a liquid polymer) 615 is dispensed from the spinneret. In one exemplary embodiment, the polymer dispensing 615 forms a plurality of polymeric fibers that extend from the center electrode to the peripheral electrode to create a layer of radially aligned fibers.

Some embodiments facilitate the creation of a concentric structure of radially aligned fibers using multiple peripheral electrodes. In one embodiment, the peripheral electrode is an internal peripheral electrode. An outer peripheral electrode defining an outer closed area larger than the inner closed area is electrically charged 620 with the first amplitude and / or polarity. The electrical charge may or may not be removed 622 from the central electrode and / or the inner peripheral electrode. The polymer is dispensed 625 from the spinneret to create an outer area of radially aligned fibers extending from the inner peripheral electrode to the outer peripheral electrode.

In addition, some embodiments facilitate the creation of a multilayer structure that includes radially aligned fibers and non-radially aligned fibers. The electrical charge 630 is removed from the peripheral electrode (s) and the center electrode. A conductive surface under the layer of radially aligned fibers is charged electrically 635 with the first amplitude and / or polarity. The polymer is dispensed 640 from the spinneret to create a layer of non-radially aligned fibers (eg, randomly oriented and / or uniaxially aligned) over the layer of radially aligned fibers.

Figure 12 is a flow chart of an example method 700 for repairing a defect in biological tissue. The defect can include a void, damage, and / or any other condition that results in decreased function of the biological tissue. In one embodiment, method 700 includes creating 705 a void in the biological tissue, and the defect is the void created. For example, the void can be created 705 through a surgical incision to provide access to an underlying tissue (eg, a tumor). In another example, the vacuum is created 705 by cutting necrotic tissue (eg, skin cells). One or more biomedical patches 710 capable of covering the defect are selected. For example, a plurality of biomedical patches 710 can be selected for a large and / or complex (eg, irregularly shaped) defect. The biomedical patch includes a plurality of radially aligned polymeric fibers that extend from a center of the biomedical patch to a perimeter of the biomedical patch. For example, a biomedical patch 710 may be selected that has a diameter greater than the diameter of an approximately circular defect.

The selected biomedical patch 710 may also include non-radially aligned polymeric fibers (eg, randomly oriented and / or uniaxially aligned). For example, radially aligned fibers and non-radially aligned fibers can be configured in separate layers.

In some embodiments, the biomedical patch includes multiple radially aligned fiber areas. In one embodiment, a first set of radially aligned fibers extends from a center of the biomedical patch to a first perimeter and defines an internal area. A second set of radially aligned fibers extends from the first perimeter to a second perimeter and defines an outer area.

A substance such as a growth factor and / or a drug (eg, a chemotherapeutic drug) 715 can be applied to the biomedical patch. For example, the biomedical patch can be immersed in such a substance to allow it to occupy a cavity within the hollow fibers of the biomedical patch, dope the polymer comprising the fibers in the biomedical patch, or coat the surface of the fibers within the biomedical patch.

The biomedical patch is applied 720 to the biological tissue (eg, overlaps it) to cover at least a portion of the defect. For example, the biomedical patch can be applied 720 to dura tissue, heart tissue, and / or any biological tissue that includes a defect. In one embodiment, the perimeter of the biomedical patch extends beyond the perimeter of the defect such that the entire defect is covered by the biomedical patch. In some embodiments, the biomedical patch is attached 725 to biological tissue by a plurality of sutures, adhesives, and / or other means for attaching the biomedical patch to biological tissue. In an alternative embodiment, the biomedical patch is simply allowed to fuse with the biological tissue, for example, by adhesion of the biological cells to the biomedical patch.

After applying the biomedical patch 720 and optionally coupling it 725 to the biological tissue, the biological tissue is covered 730. In one embodiment, other tissue covering the defect (eg, dermis and / or epidermis) is repaired (eg, closed by suture). In another embodiment, one or more protective layers are applied to the biological tissue. For example, a dressing can be applied to a skin graft, with or without a protective substance, such as a gel, an ointment, and / or an antibacterial agent. In one embodiment, the protective layer includes a nanofiber structure, eg, an additional biomedical patch, as described herein.

The embodiments described herein can be used with any neurosurgical procedure involving the repair, replacement, or expansion of the dura, including, but not limited to, a transhenoidal procedure (eg, surgical removal of pituitary adenomas), various types of surgeries in the base of the skull and / or surgical removal of cranial or spinal tumors (eg, meningiomas and / or astrocytomas). In one embodiment, a biomedical patch can be applied to a bone fracture (eg, a complex fracture). In another embodiment, a biomedical patch can be applied to a defect in the skin (eg, a burn).

Likewise, such embodiments can be used to provide a dura substitute, a biomedical patch for a skin graft (eg, dermal or epidermal), a biomedical patch for tracheal repair, a frame for an artificial heart valve leaflet, a artificial mesh for the surgical repair of the gastrointestinal tract (for example, an abdominal hernia or ulcer) and an artificial mesh for the surgical repair of heart defects. For example, a cardiac biomedical patch that includes radially aligned fibers can be used to promote regeneration of cardiomyocytes. The embodiments described herein facilitate the provision of a cardiac patch of sufficient flexibility to allow movement of the biomedical patch through biological tissue (eg, cardiomyocytes).

In some embodiments, a biomedical patch is less thick than the biological tissue being repaired. As cells migrate along the radial fibers of the biomedical patch, the biological tissue regenerates.

Biomedical patches with radially aligned polymeric fibers facilitate reduction in the cost of tissue repair, improve tissue healing time, and reduce or eliminate the risk of zoonotic infection. Furthermore, such biomedical patches are relatively simple to manufacture, allowing for adaptation of shape, size and chemical composition and better availability and lack of immunogenicity. Additionally, biomedical patches with radially aligned polymeric fibers exhibit excellent handling properties due to their fabric-like composition, eliminate the need for a second surgery to harvest autologous graft tissue, and reduce the risk of contracture and adhesion compared to other products. known.

Experimental results

The dura mater is a complex fibrous membrane consisting of numerous cells and cell types, extracellular matrix proteins, and trophic factors, all of which play an important role in the colonization and duralization of artificial dural substitutes, as well as their successful implementation. of said biomedical patches in vivo. In order to assess the ability of radially aligned nanofibers to interact with natural dura, promote host cell adhesion to the graft, and enhance host cell migration throughout the graft, an ex vivo model of the graft was developed. Surgical repair of a small dural defect.

In a standard procedure, an "artificial dural defect" was introduced into a piece of dura (1 cm x 1 cm) by microsurgically sectioning a small circular hole, 7 mm in diameter, in the center of the specimen. Next, a nanofiber-based scaffold was used to repair the artificial defect by overlaying the graft on the dural sample.

Figure 13 is a schematic illustration of biological cells extending from intact dural tissue, located adjacent to the edge of a scaffold, to the central portion of the scaffold along radially aligned nanofibers. The graft covered the entire simulated dural defect, while simultaneously coming into contact with the dural tissue at the periphery of the specimen, and demonstrates the ability of native cells in intact tissue to easily adhere and migrate through the scaffolds. of nanofibers.

Figures 14A-14D are a collection of fluorescence micrographs comparing cell migration when dural tissues were grown on radially aligned nanofiber scaffolds (Figures 14A and 14C) and randomly oriented nanofibers (Figures 14B and 14D) for 4 days using a custom-made cell culture system (Figure 15). Figures 14C and 14D are enlarged views of the central portion shown in, respectively, Figures 14A and 14B. The arrow marks the center of the frame.

As shown in Figure 14A, fluorescein diacetate (FDA) stained dural fibroblasts migrated from the surrounding tissue along the radially aligned nanofibers and beyond to the center of the circular scaffold after incubation for 4 days. It was found that the cells could completely cover the entire scaffold surface in 4 days. In contrast, a void was observed after the same incubation time period for a scaffold made of random fibers (Figure 14B), indicating faster migration of native cells in the radially aligned nanofiber scaffolds than their counterparts. random. It is clear that the scaffold made of radially aligned nanofibers (shown in Figures 14A and 14C) was completely populated with dural cells that had migrated from the edges of the adjacent dural tissue. In contrast, an acellular region is clearly visible in the center of the scaffold formed by the randomly oriented nanofibers after the same incubation time, indicating that cellular infiltration was incomplete and occurred at a slower rate.

In order to further investigate the effect of fiber alignment and subsequent modification of the nanofiber scaffold on cell migration, primary dural fibroblasts isolated from dura tissue were cultured on randomly oriented, radially aligned nanofiber scaffolds with and without fibronectin coating. Figure 15 is a schematic diagram of a custom made cell culture system designed to model tissue defect wound healing. Specifically, dural fibroblasts were selectively seeded around the periphery of a circular nanofiber scaffold, effectively forming a 7mm "simulated dural defect" in the center of the sample.

Figures 16A-16D are fluorescence micrographs showing cell morphology and distribution in scaffolds of radially aligned nanofibers (Figures 16A and 16C) and randomly oriented nanofibers (Figures 16B and 16D) with and without fibronectin coating after of a 1 day incubation. As shown in Figure 16A, many cells were able to bind to the uncoated scaffolds that include radially aligned nanofibers. In comparison, fewer cells poorly bound to scaffold without randomly oriented nanofiber coating and cell aggregations were observed (Figure 16B). The seeded cells were evenly distributed over the entire surface of the fibronectin-coated scaffold of radially aligned nanofibers, and exhibited an elongated shape parallel to the alignment axis of the nanofibers (Figure 16C). This result indicates that the fibronectin coating could improve the influence of topographic indications on the cell morphology provided by the aligned fibers. Cells were also able to adhere adequately to the fibronectin-coated scaffold of randomly oriented nanofibers and cell distribution was more uniform. than in the uncoated samples, although no elongation or cell alignment was observed (Figure 16D). The random organization of cells in the randomly oriented nanofiber scaffolds also mimics the organization of cells in scar tissue. This indicates that aligned scaffolds can help reduce scar tissue formation by promoting more regular cell organization / function.

To characterize cell motility in the scaffold, cells were stained with FDA and fluorescence images were taken at different time points. Figures 17A-17D are fluorescence micrographs showing the migration of dura fibroblasts seeded on radially aligned nanofiber fibronectin-coated scaffolds for 1 day (Figure 17A), 3 days (Figure 17B), and 7 days (Figure 17C). ). Figure 17D is an enlarged view of Figure 17C. The cells were radially aligned, replicating the alignment of the underlying fibers, as shown in Figure 17D.

The ability of dural fibroblasts to migrate and repopulate a simulated dural defect was measured as an estimate of the regenerative capacity of the surrogate at various times throughout the experiment. Figure 18 is an illustration of the determination (eg, calculation) of the area of the simulated dural defect that is still present in the framework at a given time. Figure 19 is an illustration of the area of void space as a function of incubation time. In Figure 19, "random" indicates samples with a scaffold of random fibers; "Random F" indicates samples with a fibronectin coated random fiber scaffold; "Aligned" indicates samples with a radially aligned fiber framework, and "aligned F" indicates samples with a radially aligned fiber framework coated with fibronectin. An asterisk (*) and a pound sign (#) indicate a probability p <0.05 for the samples compared to the random samples and the F random samples in the same incubation time period.

The void area decreased with increasing incubation time for all scaffolds tested due to inward migration of cells. As illustrated in Figures 17A-17D, aligned fibers can significantly improve cell migration compared to random fibers, and the fastest cell migration occurred in the fibronectin-coated radially aligned nanofiber scaffold for the first 3 days of incubation. Approximately 5 mm2 of surface area remained uncovered by cells in the uncoated random scaffolds even after incubation for 7 days. Instead, the cells covered almost the entire area of the simulated defect within the same incubation period in the other three scaffold types.

Figures 20A-20D are fluorescence micrographs showing membrane-stained living dural fibroblasts in radially aligned nanofiber scaffolds coated with fibronectin after a 1-day culture (Figure 20A), a 3-day culture ( Figure 20B), a 7-day culture (Figure 20C) and a 10-day culture (Figure 20D). Figure 20D includes an inset of a high magnification image of Figure 20D indicating that cells were radially aligned in the aligned scaffolds. Cell migration towards the center of a radially aligned nanofiber scaffold coated with fibronectin was also confirmed by images taken at preset intervals in Figures 20A-20D.

Dural tissue is mainly composed of type I collagen. Production of type I collagen from dural fibroblasts was also examined. Figures 21A-21D are fluorescence micrographs obtained by immunostaining of type I collagen with cell nuclei with 4 ', 6-diamidino-2-phenylindole (DAPI) in blue for radially aligned fiber scaffolds (Figures 21A and 21C) and randomly oriented fibers (Figures 21b and 21D). Cells in radially aligned fiber scaffolds were observed to produce comparable levels of type I collagen when compared to random fiber scaffolds, although a previous study showed that more elongated cells expressed higher type I collagen than cells. less elongated cells. Furthermore, the fibronectin coating did not have a significant influence on type I collagen production. Type I collagen was randomly targeted for the random scaffolds, resembling the extracellular composition of amorphous scar tissue, and possessed a high degree of organization for frames aligned radially, resembling healthy connective tissue.

Recent advances in the interaction between cells and biomaterials have shown that the chemical and topographic properties of the surface of materials can regulate and control the shape and function of the cell. Cell orientation, motility, adhesion, and shape can be modulated by specific surface micro- and nanotopographies. Cells can be aligned along microgrooves or similar topographic features on a surface. Fibroblasts were shown to be the most sensitive cell type compared to endothelial cells and smooth muscle cells, responding with robust alignment, elongation, and migration along these topographic features.

Simultaneously, electrospinning has been widely used to produce nanofibers for a wide variety of applications in tissue engineering, including skin grafts, artificial blood vessels, nerve repair, and others. However, previous studies were limited to the use of scaffolds made of randomly and uniaxially aligned nanofibers. Scaffolds composed of uniaxially aligned nanofibers are impractical for wound healing applications because wounds are often irregular in shape. The work described herein demonstrated for the first time the fabrication of a new type of frames made of radially aligned nanofibers. This new type of scaffold can guide dural fibroblasts extending in the direction of fiber alignment and direct cell motility toward the center of the scaffold, resulting in faster cell migration and infiltration in comparison. with scaffolds composed of randomly oriented nanofibers.

Furthermore, uniaxially aligned nanofiber scaffolds cannot match such capacity, as they can only guide cell migration in one direction. It was reported that the control of cell orientation or morphology by topography, the so-called "contact guide", could allow the organization of the extracellular matrix. For most injuries, repair results in previously functional tissue becoming a disorganized amalgam of cells (eg, fibroblasts) and an extracellular matrix (eg, collagen fibers) known as a scar. Highly organized cells and extracellular matrix are required for proper tissue regeneration and function, which is typically very different from repair of scar tissue. It has been shown in the present work that extracellular matrix type I collagen in radially aligned nanofiber scaffolds showed a high degree of organization, indicating that radially aligned nanofiber scaffolds may reduce the possibility of scar tissue formation afterwards. of wound healing.

A dura substitute must be safe, effective, easy to handle, impervious to water, and easy to integrate into the surrounding tissue to form new tissue similar to native tissue. Likewise, it must avoid harmful reactions caused by foreign bodies, be free from any potential risk of infection, have mechanical properties similar to those of natural dura mater -particularly with respect to its flexibility and resistance-, be stable and / or storable, and be available for immediate use. In the present work, the biodegradable polymer polycaprolactone (PCL) was chosen as the material for the dural substitute, since PCL has some advantages compared to other bioabsorbable polyesters. Heterogeneous degradation of PGA and poly (L-lactic acid) (PLLA) could result in a sudden increase in degradation products, resulting in acidic conditions and toxic reactions in the surrounding tissue. The degradation of PCL is slower and produces less acidic degradation products and has been studied as a wound dressing material since the 1970s.

In order to obtain watertight properties, the radially aligned nanofiber scaffold can be combined with a nonwoven mat to form two-layer or even multi-layer substitutes. Simultaneously, antibiotics can be easily encapsulated within the nanofibers to further reduce the inflammatory response, improve wound healing, and prevent adhesion after surgery. Alternatively, PCL can be blended with other polymers to further enhance its biocompatibility, as well as its mechanical, physical, and chemical properties. In addition, extracellular proteins and / or growth factors can be immobilized on the surface of nanofibers using various surface modification approaches to enhance cell adhesion. Current work demonstrates the effect of fibronectin coating on PCL nanofibers through electrostatic interaction on dural fibroblast adhesion and motility. The results presented herein demonstrate that fibronectin coating improved dural fibroblast adhesion and enhanced cell migration in randomly oriented nanofiber scaffolds. In contrast, coating had a marginal contribution to cell motility in radially aligned nanofiber scaffolds, compared to uncoated scaffolds, indicating the predominant role played by nanofiber alignment and the resulting surface topography.

In summary, the present describes the manufacture of a new type of electrospun nanofiber scaffold that includes radially aligned fibers and the possible application of such structures as dural substitutes. Dural fibroblasts grown on radially aligned nanofiber scaffolds elongated parallel to the axis of the fibers, and cell migration toward the center of the scaffold accelerated along with the development of a regular type I collagen-like extracellular matrix configuration, which potentially promotes rapid regeneration and neodura formation. Taken together, these results indicate that radially aligned nanofibers have great potential as an artificial dural substitute, may offer an alternative in the repair of dural defects, and, furthermore, occupy a unique and desirable place within the neurosurgical community.

Additional experimental results

In a common PCL nanofiber electrospinning procedure (Mw = 65 kDa, Sigma-Aldrich), a 20% (w / v) solution of PCL in a mixture of dichloromethane (DCM) and N, N-dimethylformamide ( DMF) (Fisher Chemical) with a volume ratio of 8: 2. The fibers were spun at 10-17 kV with a feed rate of 0.5 ml / hr onwards, along with a 23 gauge needle as the spinneret. A piece of aluminum foil was used as a collector to obtain random nanofiber scaffolds. The radially aligned nanofiber scaffolds were fabricated using a collector consisting of a ring electrode (eg, a metal ring) and a pointed electrode (eg, a sharp needle). The electrospun PCL nanofibers were coated with fibronectin (Millipore, Temecular, California) as follows. Electrospun fiber scaffolds were sterilized by soaking in 70% ethanol overnight and washed three times with phosphate buffered saline (PBS). The scaffolds were then soaked in 0.1% poly-L-lysine (PLL) solution (Sigma-Aldrich) for one hour at room temperature, and then washed with PBS buffer (Invitrogen) three times.

Subsequently, the samples were immersed in a fibronectin solution (26 µl 50 µg / µl of fibronectin solution diluted with 5 ml of PBS buffer) at 4 ° C overnight. Prior to cell seeding, the fibronectin solution was removed and the nanofiber scaffolds were rinsed with PBS buffer.

The PCL nanofiber scaffolds were sputter coated with gold prior to imaging with a scanning electron microscope (Nova 200 NanoLab, FEI, Oregon, USA) with an acceleration voltage of 15 kV. Samples prepared for use in cell cultures were inserted into a 24-well cell culture treated polystyrene (TCPS) culture plate and sterilized by soaking the scaffolds in 70% ethanol.

Fibroblasts were isolated from explanted dura sections. Specifically, a 2.0 cm x 1.5 cm section of dura was removed by sharp dissection, which was washed three times with cold PBS. Dural fibroblasts were then isolated by softening chopped dura mater three times in 4 ml of Hank's Balanced Salt Solution (HBSS) containing 0.05% Trypsin and 0.04% EDTA (Sigma-Aldrich, St. Louis, Missouri). After softening, the collected supernatant was centrifuged and the dural cell pellet isolated and resuspended in Dulbecco's Modified Eagle's Medium (DMEM) supplemented with 10% bovine serum and 1% penicillin and streptomycin. Dura cells obtained in this way were plated in 75 cm2 flasks and expanded (transferred to a new culture medium no more than 5 times).

Large continuous pieces of dura were placed in cold PBS and microsurgically cut into 1 cm x 1 cm sections. An artificial defect was then introduced into each section of the dura by microsurgically cutting a small circular hole, 7 mm in diameter, in the center of the section. Dura sections were then introduced into individual wells of 6-well culture plates containing 4 ml DEMEM supplemented with 10% bovine serum and 1% penicillin and streptomycin. Subsequently, radially and randomly aligned nanofiber scaffolds 1 cm in diameter were used to repair the artificial defects by overlaying the graft on the dural sample. Nanofiber scaffolds were placed over the dura mater, in such a way that the graft covered the entire defect and at the same time came into contact with the dural tissue at the periphery of the specimen. The nanofiber scaffolds were held in this position throughout the experiment by placing a sterile metal ring over the scaffold and dura. After 4 days of culture, the cells were stained with FDA in a green color and images were obtained with a fluorescence microscope. Fluorescence images were taken using a 12-bit QICAM Fast Cooled Mono camera (Q Imaging, Burnaby, British Columbia, Canada) coupled to an Olympus microscope with OCapture 2.90.1 (Olympus, Tokyo, Japan). Similarly, about 1 x 105 dural fibroblast cells were seeded on the periphery of the nanofiber scaffolds using the custom-made culture system shown in Figure 15. After different time periods, the cells were stained with FDA. in green color and images were obtained with a fluorescence microscope. The total surface area of the nanofiber scaffold devoid of migrating cells was then calculated using Image J software (National Institute of Health).

Live cells were labeled with a membrane stain using VYBRANT DiO cell labeling solution (Invitrogen), following the manufacturer's instructions, and then imaged on days 1, 3, 7, and 10.

Type I collagen production by dural fibroblasts in fiber scaffolds was evaluated by immunohistochemistry. On day 7, cells were rinsed with PBS and fixed with 3.7% formalin for 1 hr (N = 4). Cells were permeabilized using 0.1% Triton X-100 (Invitrogen) in PBS for 20 min, followed by blocking in PBS containing 5% normal goat serum (NGS) for 30 min. Monoclonal antibodies to type I collagen (1:20 dilution) were obtained from EMD Chemicals (Calbiochem, San Diego, California). Cells were washed three times with PBS containing 2% FBS. Secondary antibody Gt x Rb IgG Fluor (Chemicon, Temecula, California) (1: 200 dilution) was applied for 1 hour at room temperature. Fluorescence images were taken using a 12-bit QICAM Fast Cooled Mono camera (Q Imaging, Burnaby, British Columbia, Canada) coupled to an Olympus microscope with OCapture 2.90.1 (Olympus, Tokyo, Japan).

Mean values and standard deviation were obtained. Comparative analyzes were carried out using Tukey's post hoc test by analysis of variance at a 95% confidence level.

As a secondary study, an ex vivo model of the surgical repair of a small dural defect was developed. Large pieces of healthy dura (3 cm x 3 cm) were placed in cold supplemented Dulbecco's Modified Medium (DMEM) and cut into smaller pieces (1 cm x 1 cm) microsurgically. Artificial defects were introduced into the dura pieces by microsurgically cutting small circular holes, 6-8 mm in diameter, in the center of the specimens. The radially aligned nanofiber scaffolds, randomly oriented nanofiber scaffolds, and DURA MATRIX collagen scaffolds (1 cm x 1 cm) were then used to repair the artificial defects by overlaying the graft on the dural sample, such that the graft covered the entire defect, while coming into contact with the dural tissue in the periphery of the sample.

The dural / dural substitute pools were cultured in vitro in supplemented DMEM for a period of four days. At the end point of this period, light and fluorescence microscopy was used to assess the ability Regenerative surrogate, defined as the ability of the dural cells to migrate to the artificial surrogate and repopulate the acellular region of the dural surrogate within the artificial defect.

The results demonstrated that native cells present in intact dura (mainly dural fibroblasts) readily migrated to adjacent polymeric nanofiber dural substitutes in high concentrations within 24 to 48 hours after coming into contact with explanted dura fragments. Migration of dural cells to reference collagen matrices occurred in a similar period, although slightly lower concentrations of dural cells migrating to collagen matrices were observed when compared to nanofiber dural substitutes. This observation indicates that nanofiber dural substitutes readily adhere to native dural tissue, an important quality with regard to intraoperative material handling and / or placement, and that nanofiber dural substitutes provide an ideal substrate for adhesion of dural fibroblasts.

Further examination of the various dural substitutes after four days of culture revealed that dural migration of fibroblasts to the central acellular region of the material proceeded significantly more rapidly in radially aligned nanofiber substitutes than in randomly oriented nanofiber substitutes or matrices. collagen. This discovery was evidenced by the fact that after four days of culture, a prominent acellular region ("empty space") remained in the samples of both the random nanofiber surrogate and the collagen matrix.

Instead, the radially aligned nanofiber material samples, examined at the same time point, had been completely populated with dural cells that had migrated from the edges of the adjacent dural tissue. Indeed, the radially aligned nanofiber substitutes were able to cause significantly faster "healing" of this simulated dural defect than both randomly oriented materials. High-magnification views of the dural substitutes within this ex vivo culture further demonstrated the ability of radially aligned nanofiber materials to align and direct native migratory dural cells, a similar result to the previous study conducted using pre-seeded dural fibroblasts. . Specifically, it was observed that the dural cells align and extend parallel to the individual nanofibers within the artificial substrate, and also deposit organized extracellular matrix proteins (specifically type I collagen) on the aligned nanofiber materials. This observation indicates that topographic indications presented by aligned nanofiber substitutes are capable of organizing and directing native dural cells that migrate from intact dura and may enhance the ability of these migratory cells to deposit extracellular matrix proteins necessary to heal and repair the cells. dural defects.

The results of this secondary study demonstrate that nanofiber dural substitutes not only provide a favorable scaffold for dural cell adhesion and migration, but easily promote dural cell inward growth from the entire intact dura. The ability of nanofiber materials to intimately interconnect with the intact dura and facilitate rapid cell population of the polymeric scaffold clearly indicates that this material can function exceptionally well as an artificial graft in the surgical repair of complex dural defects. Additionally, dural substitutes constructed from radially aligned nanofibers were shown to promote faster "healing" of simulated dural defects than randomly oriented materials, indicating that aligned nanofiber scaffolds that impart nanoscale topographic features may represent a significant technological advance over benchmark clinical collagen matrices.

Although the experiments described herein were of limited duration, the results of these experiments indicate that biomedical patches that include radially aligned fibers are viable for use in tissue repair over longer periods of time. For example, it is anticipated that the accelerated cell growth observed would continue until biological tissue at the site of a defect is fully regenerated and / or until degradation of the biomedical patch is complete.

In vivo experimental results

In vivo experimentation was carried out by imposing a 12 millimeter diameter dural defect in a native porcine dura. The defect was repaired with a collagen dural substitute, a single-layer dural substitute with randomly oriented nanofibers, and a two-layer dural substitute with a radially aligned nanofiber layer fused with a second layer of randomly oriented nanofibers by layer stacking. per layer (eg, as described above with reference to Figure 8). In a control group, the defect was not repaired.

Figure 22 is a graph 2200 illustrating the thickness of the regenerated dura at the center of repaired dural defects over time. In graph 2200, the "y" axis 2205 represents the total thickness of the regenerated dura, including both regenerative tissue and integrated dural substitute material, at the center of a dural defect. Samples without a dural substitute (control samples) or with a collagen dural substitute, a single-layer randomly oriented nanofiber dural substitute, and a two-layer radially aligned nanofiber dural substitute are grouped by elapsed time on the x-axis. ”2210.

Figure 23 is a graph 2300 illustrating the content of regenerative collagenous tissue over time. In graph 2300, the "y" axis 2305 represents the percentage of regenerated dura that is composed of regenerative collagenous tissue. Samples with a collagen dural substitute, a single-layer randomly oriented nanofiber dural substitute, and a two-layer radially aligned nanofiber dural substitute are grouped by elapsed time on an "x" axis 2310.

Electrode Arrays

In some embodiments, a collector includes a plurality of electrodes that at least partially circumscribe an area and a second electrode located within that area. The electrodes can be configured in a matrix, such as a grid and / or other polygonal pattern, and a polymer deposited on the electrodes can form fibers that run between the collector electrodes, such that the fibers define the sides of a plurality of polygons, with the electrodes located at the vertices of the polygons. In some embodiments, the structure created by said fibers can be used to create a cellular microarray, for example, by seeding the structure with cells and incubating the cells to promote the propagation of the cells throughout the structure.

Cell microarrays can provide powerful experimental tools for high throughput screening, useful in a number of applications ranging from drug discovery and toxicology to stem cell research and tissue engineering. For example, cell microarrays may represent an effective means of fabricating ordered neural networks useful for studying synapse formation and neuronal plasticity in vitro. At least some known techniques for the fabrication of neuronal microarrays have concentrated on the use of spatial patterns of cell-adhesive and / or cell-repellent materials and agents. Unfortunately, such manufacturing techniques can be time-consuming and expensive, and involve the use of sophisticated instruments (for example, photolithography, soft lithography, contact printing, microfluidic technique, nano printing, and inkjet printing. ink).

Electrospinning is capable of producing one-dimensional fibers with diameters ranging from several nanometers to several microns. The high surface-to-volume ratio and nanoscale morphology of electrospun nanofibers may indicate that these materials effectively mimic the architecture of the extracellular matrix (ECM). As a result, electrospun nanofiber materials have been used in a wide variety of biomedical applications. Electrospun nanofibers can be randomly deposited on a conductive collector and / or aligned in uniaxial arrays by manipulating an electric field and / or applying mechanical forces.

The embodiments described herein facilitate the production of a complex cell microarray using electrospun nanofibers. In exemplary embodiments, a collector with an electrode array is used to fabricate electrospun nanofiber scaffolds that include a complex array architecture and numerous multiple wells. Said scaffold may be useful at least for: (i) the formation of cell microarrays; and (ii) the formation of neural networks. The use of presented complex nanofiber arrays can facilitate the creation of advanced substrates useful in neural engineering applications and cell arrays useful in drug pre-screening and biosensing applications.

Figure 24 is a diagram illustrating a perspective view of an example electrospinning system 2400 for producing a polygonally aligned fiber structure using an electrode array. System 2400 is similar to System 100 (shown in Figure 1) in structure and operation. A collector 2405 includes a plurality of first electrodes 2410, which may be referred to as peripheral electrodes. The first electrodes 2410 define and / or at least partially circumscribe an area 2415, such as a polygon. As illustrated in Figure 24, the area 2415 defined by the first electrodes 2410 is a hexagon. A second electrode 2420, which may be referred to as an internal electrode, is located within (eg, approximately in the center) of area 2415, such that the first electrodes 2410 surround the second electrode 2420. In example embodiments, the first electrodes 2410 and the second electrodes are metallic spheres (eg stainless steel) having a diameter between 0.5mm and 5.0mm (eg 1.0mm or 2.0mm).

The 2400 system also includes a row 120 and is configured to create an electrical potential between the collector 2405 and the row 120, as previously described with reference to Figure 1. In exemplary embodiments, the peripheral electrodes 2410 and the inner electrode 2420 are electrically coupled to a power source 130 through conductor 135, and row 120 is coupled to power source 130 through conductor 145. Power source 130 is configured to charge peripheral electrodes 2410 with a first amplitude and / or polarity through lead 135, and to charge row 120 with a second amplitude and / or polarity, opposite the first polarity, through lead 145.

In the embodiment illustrated in Figure 24, peripheral electrodes 2410 and inner electrode 2420 are metallic spheres or balls (eg, stainless steel), which may be referred to as "microspheres", configured in a hexagonal pattern. In some embodiments, the circular closed area 125 may have a diameter of between 1 cm and 20 cm. In other embodiments, the peripheral electrodes 2410 and the inner electrode 2420 can be of any shape and / or may be configured in any appropriate pattern for use with the methods described herein. For example, peripheral electrodes 2410 and inner electrode 2420 can be shafts, rods, domes, and / or ridges. Additionally, peripheral electrodes 2410 and inner electrode 2420 may be configured in an octagonal, pentagonal, and / or square pattern, for example, although other polygonal and non-polygonal, regular and / or irregular configurations are also envisioned.

In one embodiment, area 2415 defines a horizontal plane 2425. Row 120 is aligned with inner electrode 2420 and is offset vertically from horizontal plane 2425 by a variable distance. For example, row 120 may be vertically offset from horizontal plane 2425 by a distance of 1 cm to 100 cm. In exemplary embodiments, the inner electrode 2420 and / or peripheral electrodes 2410 include a rounded (eg, convex) surface, such as the surface of the metal spheres shown in Figure 24, oriented toward the horizontal plane 2425.

As previously described with respect to Figure 1, spinneret 120 is configured to dispense polymer 140 while spinneret 120 is electrically charged with the second amplitude and / or polarity, and peripheral electrodes 2410 and inner electrode 2420 are electrically charged with the first amplitude and / or polarity. Row 120 dispenses polymer 140 as stream 160. Stream 160 has a diameter approximately equal to the diameter of the opening of row 120. Stream 160 descends toward manifold 2405. For example, stream 160 may drop down low influence of gravity and / or may be drawn downward by a charged conductive surface 162 located below collector 2405. For example, conductive surface 162 may be electrically coupled to conductor 135 and charged with the same amplitude and / or polarity as peripheral electrodes 2410 and center electrode 2420. As flow 160 descends and is deposited in collector 2405, polymer 140 forms one or more solid polymeric fibers 2430 that extend from inner electrode 2420 to peripheral electrode 2410 and / or between peripheral electrodes 2410.

In some embodiments, collector 2405 includes peripheral electrodes 2410 that define a plurality of areas 2415. For example, peripheral electrodes 2410 immediately surrounding inner electrode 2420 can be considered inner peripheral electrodes, and a plurality of outer peripheral electrodes 2435 can surround the inner peripheral electrodes 2410, such that the inner peripheral electrodes 2410 are nested within the outer peripheral electrodes 2435. The collector 2405 may include any number of nested sets of peripheral electrodes. Although the collector 2405 includes electrodes in a compact configuration (for example, with electrodes that are in contact with each other), it is envisaged that the electrodes may be separated from each other by an electrode distance, which may be constant throughout the collector or may vary between different pairs of electrodes.

Furthermore, in some embodiments, a collector may include electrodes that define a plurality of partially overlapping areas in a modular manner. Figure 25 is a diagram illustrating a perspective view of an example modular electrospun collector 2500. Collector 2500 includes first electrodes 2505 surrounding a second electrode 2510. First electrodes 2505 define a first hexagonal area 2515 With respect to the first hexagonal area 2515, the second electrode 2510 can be considered an internal electrode, and the first electrodes 2505 can be considered peripheral electrodes.

The collector 2500 also includes a plurality of third electrodes 2520 that are located outside of the first hexagonal area 2515. The third electrodes 2520, second electrode 2510, and a subset of first electrodes 2505 define a second hexagonal area 2525 that partially overlaps the first hexagonal area 2515. One of the first electrodes 2505 (for example, a peripheral electrode with respect to the first hexagonal area 2515) is located within the second hexagonal area 2525. With respect to the second hexagonal area 2525, this first electrode 2505 can be considered an internal electrode. The third electrodes 2520, the subset of the first electrodes 2505, and the second electrode 2510 can be considered peripheral electrodes. Although the electrodes defining two partially overlapping areas are illustrated in Figure 25, it is envisaged that the modular nature of collector 2500 facilitates the inclusion of any number of electrodes defining any number of areas, such that collector 2500 can be extended in one or more directions by adding electrodes to the perimeter of the collector 2500.

As previously described with reference to system 2400 (shown in Figure 24), collector 2500 (eg, first electrodes 2505, second electrode 2510, and third electrodes 2520) is configured to be electrically charged with an amplitude and / or polarity opposite to the amplitude and / or polarity with which the row 120 is electrically charged. When these components are thus charged, a polymer dispensed from spinneret 120 can form fibers that extend between electrodes (eg, first electrodes 2505, second electrode 2510, and / or third electrodes 2520) of collector 2500.

Figure 26 is a 2600 diagram illustrating an electric field generated by an electrospinning system such as the 2400 electrospinning system (shown in Figure 24). Diagram 2600 shows a two-dimensional cross-sectional view of electric field intensity vectors between a row 120 and a plurality of electrodes 2605.

The electric field vectors near the surface of the electrodes 2605 are oriented perpendicular to the surface of the electrodes 2605. The electric field vectors between two neighboring electrodes are divided into two main flows, pointing toward the centers of the two adjacent electrodes 2605 Accordingly, the fibers deposited on the surface of the electrodes 2605 can be randomly distributed, while the fibers deposited in the area between two neighboring electrodes 2605 can be uniaxially aligned between these two adjacent electrodes 2605.

Figures 27A-27F are microscopy images of a 2705 nanofiber membrane produced using a collector with an electrode array, such as collector 2405 (shown in Figure 24). For example, membrane 2705 can be produced using a matrix of stainless steel spheres. Figure 27A is a light microscopy image of a membrane 2705. Figure 27A includes an inset 2710 illustrating a magnification of the membrane 2705 with a light source on the right side of the image. The shadows in box 2710 indicate wells within membrane 2705, the positions of which correspond to the positions of the electrodes on the collector.

Figure 27B is a scanning electron microscopy (SEM) image of membrane 2705 illustrating the complex ordered architecture composed of hexagonally arranged wells 2715 connected with uniaxially aligned fiber arrays 2720. The depth of wells formed by depositing electrospun nanofibers on 1.0 mm and 2.0 mm diameter compact stainless steel microspheres were approximately 200 microns (pm) and 400 pm, respectively. Such wells can be referred to as "microwells".

Figures 27C-27F are enlargements of corresponding areas within Figure 27B. Figure 27C indicates that the fibers deposited on the surface of the microsphere electrodes were randomly distributed. Figure 27D shows that the fibers at the interface between an electrode surface and an electrode gap transitioned from a random orientation to an aligned orientation. Figure 27E indicates that the fibers deposited along the axis connecting the centers of two adjacent electrodes were uniaxially aligned parallel to that axis. Figure 27F shows that the fiber density was significantly lower between neighboring spheres and away from the axes connecting the centers of adjacent spheres than in other areas (for example, those shown in Figures 27c -27E), and that the deposited fibers in this area they were randomly oriented.

In some embodiments, a fiber membrane, such as membrane 2705, can be combined with other membranes. For example, a membrane with a plurality of wells interconnected by uniaxially aligned fibers can be used as a layer within a multi-layer structure, as described above with reference to Figure 8. Additionally or alternatively, different types of collectors, for example, using an electrode array collector as an internal collector (for example, corresponding to a center of a biomedical patch), and using a ring-type collector (for example, as shown in Figure 1) as an outer manifold that surrounds the inner manifold.

Experimental results

The fiber membranes or "scaffolds" produced by an electrode array collector, as described above, were evaluated for use as substrates to generate cell microarrays. Cells were selectively seeded on the scaffold surface by placing a small amount of media, which contained a specific number of cells, on the microwells present within the nanofiber arrays.

Figures 28A-28D are microscopy images illustrating cell growth on a membrane such as membrane 2705 (shown in Figures 27A-27F). Figure 28A is a light microscopy image illustrating that cell-containing droplets 2805 can be placed within wells of a fiber membrane. In addition, the hydrophobic fibers can facilitate the maintenance of these drops for more than two hours. Cells adherent to the nanofiber arrays were found to be weakly bound after two hours and easily removed using a PBS buffer, indicating rapid and reversible attachment of cells within the microarrays. Cells adherent to the nanofiber arrays after twenty-four hours were stained with fluorescein diacetate (FDA) in green to identify living cells.

Figures 28B-28D are fluorescence microscopy images illustrating cell microarrays. Live MG-63 cells were stained with fluorescein diacetate and are shown in Figures 28B-28D as light areas against a dark background.

A matrix of cells selectively adhered to microwells within the nanofiber membrane is shown in Figure 28B. Each well within the scaffold was observed to contain approximately 45 cells, while very few cells were observed outside the microwells within the fiber membrane. The average number of adherent cells in each microwell was easily manipulated by controlling the density of the cells present within the seed drops.

Figure 28C demonstrates cell microarrays seeded with a greater number of cells (approximately 150 cells per well) than are used in the arrays shown in Figure 28B. Despite the increase in the cell concentrations, the cells remained highly confined in the wells of the nanofiber scaffold. The same cell microarray illustrated in Figure 28C is shown in Figure 28D after incubation for three days. Comparison of Figure 28D and Figure 28C demonstrates that the seeded cells were capable of proliferating and migrating on the surface of the nanofiber scaffolds, although they generally remained physically confined within the wells of the cell microarray.

To examine the potential of these unique nanofiber scaffolds as effective substrates for neuronal engineering applications, dorsal root ganglia (DRG) were seeded on polylysine and laminin functionalized fiber membranes and incubated for 6 days. The resulting neurite fields protruding from the DRGs were stained with antineurofilament 200 to visualize neurite spread along the underlying nanofiber scaffold.

Figures 29A and 29B are microscopy images illustrating neurite propagation in a membrane such as membrane 2705 (shown in Figures 27A-27F). Figure 29A is an overlay of a light microscopy image and a fluorescence microscopy image illustrating that neurites emanated from a DRG main body located in the center of Figure 29A and formed an appreciable neural network after 6 days of culture. Neurites were observed to grow along the long uniaxially aligned nanofiber axes and reach neighboring microwells, effectively reproducing the geometry of the underlying nanofiber architecture.

Figure 29B is an overlay of a light microscopy image and a fluorescence microscopy image adjacent to the area shown in Figure 29A. Figure 29B demonstrates that neurites can continue to grow along the nanofiber uniaxial alignment direction after reaching neighboring wells and navigating to neighboring wells along the fiber alignment in various directions. Subsequently, it was observed that the neurites that extended to the adjacent microwells were divided into five groups following the aligned fiber arrays that connected to a secondary set of adjacent wells, further indicating the scaffold's ability to form a complex neural network in vitro. .

Figures 30A and 30B are overlays of a light microscopy image and a fluorescent microscopy image illustrating the formation of neural networks from embryoid bodies in a membrane, such as the membrane shown in Figures 27A-27F. Embryonic stem cells (ES), cultured to aggregate and form embryoid bodies (EB) using the 4- / 4 + protocol, were seeded in electrospun nanofiber scaffolds such as those shown. shown in Figures 27A-27F, and incubated with supplement B27 to induce neuronal differentiation. Immunostaining with Tuj1, a neuronal marker, was performed after incubation for 14 days in order to examine the ability of the underlying nanofiber scaffolds to promote neuronal differentiation in vitro.

Figures 30A and 30B demonstrate the ability of embryoid bodies to form neural networks on nanofiber membrane substrates. In one case, an embryoid body was confined within one of the microwells, while neurites spread peripherally along the underlying fiber pattern, as shown in Figure 30A. Neurites extending from the cultured embryoid bodies were similarly aligned in the uniaxial part of the scaffold where the fibers were highly organized. Upon reaching the area of adjacent wells, the neurites arranged randomly as a result of the random orientation of the underlying fibers.

In another case, embryoid bodies were seeded in areas of nanofibers aligned uniaxially within the nanofiber array, as shown in Figure 30B. Again, the neurites spread out along the alignment direction of the fibers and, upon reaching the closest well, exhibited a disorderly organization. In particular, when the neurites spread through the microwell area, their uniaxial alignment was restored, parallel to the alignment of the underlying fibers. Taken together, these results indicate that the nanofiber architectures described herein represent a simple and efficient means of developing complex neural networks from primary neurons or embryonic stem cells.

Experimental procedure

The electrospinning system used to fabricate and collect aligned nanofibers was similar to the 2400 system (shown in Figure 24). The polymer solution used for electrospinning contained 20% PCL (w / v) in a mixed solvent of dichloromethane (DCM) and dimethylformide (DMF) with a volume ratio of 80:20. The manifold included sets of stainless steel microspheres with diameters of 1 mm and 2 mm, respectively. The fiber membranes were transferred to culture plates and then fixed by medical grade silicone adhesive. The PCL fibers were sputter coated with gold prior to scanning electron microscope imaging at an accelerating voltage of 15 kV.

For dorsal root ganglia (DRG) culture and immunostaining, DRGs were dissected from the thoracic region of day 8 embryonic chicks (E8, stage HH35-36) and harvested in Hank's buffered saline (HBSS). prior to plating. DRGs were seeded into the fiber architectures and incubated for 6 days in modified neurobasal media (NB) containing 1% ABAM, 1% supplement of N-2 (Invitrogen) and 30 ng / ml of beta nerve growth factors (B-NGF) (R&D Systems, Minneapolis, Minnesota). After incubation for 6 days, the DRGs were immunostained with the anti-neurofilament marker 200 (Sigma-Aldrich). Briefly, DRGs were fixed in 3.7% formaldehyde for 45 minutes and permeabilized with 0.1% Triton X-100 for 30 minutes. Samples were blocked in PBS containing 2.5% bovine serum albumin (BSA) (Sigma-Aldrich) for 1 hour. Anti-NF 200 diluted with PBS containing 1.5% BSA was applied to cells overnight at 4 ° C. Next, a secondary antibody, AlexaFluor 488 anti-mouse (goat) IgG (1: 200, Invitrogen) was applied for 1 hour at room temperature. After staining, fluorescence images were captured.

For embryoid body formation and immunostaining, embryoid bodies were seeded into fiber architectures and incubated with basal neuronal media containing the B27 supplement. After 14 days, immunohistochemistry was performed to visualize the spatial distribution of neurites in accordance with our previous study.

The MG-63 cell line was used to demonstrate cell microarray formation. Cells were cultured in alpha minimal essential medium (a-MEM, Invitrogen, Grand Island, New York), supplemented with 10 % fetal bovine serum (FBS, Invitrogen) and 1% antibiotics (containing penicillin and streptomycin, Invitrogen). The medium was changed every other day and the cultures were incubated at 37 ° C in a humidified atmosphere containing 5% CO2. A certain number of cells were seeded in each well of the scaffolds by placing small drops in the wells. After incubation for 2 hours, the scaffolds were washed with culture medium to remove loosely bound cells. The living cells were then stained with fluorescein diacetate (FDA) after incubation for 24 hours and images were taken under a fluorescence microscope.

Additional electrode array configurations

In addition to the concrete examples of electrode arrays described above with reference to experimental results, it is envisioned that nanofiber structures, such as those described herein, can be produced with several different electrode arrays. Figures 31A-31D are scanning electron microscopy images illustrating membranes produced using a variety of electrode arrays.

Figure 31A illustrates a fiber membrane made using a manifold comprised of hexagonal arrays of stainless steel spheres. Illustrated in Figure 31B is a fiber membrane made using a manifold composed of hexagonal arrays of stainless steel spheres having a larger diameter than the stainless steel spheres used to produce the membrane shown in Figure 31A.

Other non-hexagonal array configurations can also be used with the electrodes to achieve different geometries. Figure 31C shows a fiber membrane made using a manifold composed of a compact square matrix with stainless steel spheres. Figure 31D shows a fiber membrane produced using a manifold composed of square arrays of stainless steel microspheres with a gradual increase in the distance between electrodes in one direction. The fiber membranes were not removed from the collectors during SEM imaging and can be easily removed (eg, peeled off) from the collectors when required.

Figure 32 is a diagram of a collector 3200 with peripheral electrodes 3205 partially circumscribing an area 3210. The collector 3200 also includes an internal electrode 3215. The peripheral electrodes 3205 and internal electrode 3215 define a portion 3220 of area 3210. In examples In embodiment, peripheral electrodes 3205 are located at a perimeter 3225 of area 3210.

In the embodiment shown in Figure 32, area 3210 is shown as an ellipse (eg, a circle), and portion 3220 is shown as a sector of the ellipse. It is envisaged that the area 3210 can be of any geometric or non-geometric shape, such as an ellipse, a polygon, an oval, a rectangle, a square, a triangle and / or any rectilinear or curvilinear shape, and that the part 3220 it can be any part of that shape.

The electrode matrix fiber structures described herein allow the formation of "dimpled" structures within a fiber membrane. Accordingly, the production of such membranes represents a significant advance as the disclosed fiber membranes possess multiple microwells arranged in varying ordered geometries. Furthermore, these structures possess unique three-dimensional microwells capable of physically confining the seeded cells on the scaffold surface and facilitating the fabrication of cell microarrays. Compared to known approaches to microarray fabrication, the use of fiber membranes can be a simpler and less expensive technique to form complex cell microarrays for in vitro and in vivo use. Furthermore, the previously described experimental results demonstrate that neurites at the well site were randomly distributed, and that neurites were able to bond well to well along intermediate aligned fibers. A neural network developed using this structure could be used for high-throughput applications in neurotoxicology and developmental neurobiology.

To facilitate understanding of this invention, a number of terms are defined below. The terms defined herein have the usual meaning they would have to a person skilled in the art in the areas relevant to embodiments of the invention. Articles such as "a", "the", "the", "the" and "the" do not refer only to a singular entity, but include the general class of which a specific example can be used as a title. illustrative.

Claims (12)

1. A method of producing a fiber structure, the method comprising: electrically charging one or more first electrodes defining an area with a first polarity; electrically charging a second electrode located within the area defined with the first polarity; electrically charging a row with a second polarity opposite the first polarity, wherein the row is aligned with the second electrode; Y dispensing a polymer from the spinneret to create a fiber structure extending from the second electrode to the first electrode (s). The method of claim 1, wherein the dispensing of the polymer comprises forming a plurality of polymeric fibers extending from the second electrode to the first electrode to create a layer of radially aligned fibers. The method of claim 2, further comprising, after creating the layer of radially aligned fibers: removing the electrical charge from the first electrode and the second electrode; electrically charging a conductive surface beneath the layer of fibers radially aligned with the first polarity; Y dispensing the polymer from the spinneret to create a layer of non-radially aligned fibers over the layer of radially aligned fibers. The method of claim 2, wherein said one or more first electrodes are one or more internal electrodes defining an internal area, and the layer of radially aligned fibers is an internal area of radially aligned fibers, and this method comprises, what's more: electrically charging one or more external electrodes defining an external area larger than the internal area with the first polarity; Y dispensing the polymer from the spinneret to create an outer area of radially aligned fibers extending from the inner electrode to the outer electrode. The method of claim 1, wherein the dispensing of the polymer comprises dispensing a liquid polymer to the second electrode. The method of claim 1, wherein charging said one or more first electrodes comprises charging a circular electrode surrounding the second electrode. The method of claim 1, further comprising applying a mask between the spinneret and said one or more first electrodes while dispensing the polymer. 8. An apparatus for producing a structure including a plurality of fibers, the apparatus comprising: a plurality of first electrodes that at least partially circumscribe an area; a second electrode located within the area and electrically coupled to the first electrodes; Y a power source electrically coupled to the first electrodes and the second electrode, wherein the power source is configured to electrically charge the first electrodes and the second electrode with a first polarity, wherein when a row aligned with the second electrode and charged with a second polarity opposite the first polarity dispenses a polymer, the dispensed polymer forms a plurality of fibers extending from the second electrode to the first electrodes. The apparatus of claim 8, wherein the first electrodes surround the second electrode. The apparatus of claim 8, wherein the first electrodes define a polygon, and the second electrode is located within the polygon. The apparatus of claim 8, wherein the first electrodes and the second electrode define a sector of an ellipse. 12. The apparatus of claim 8, wherein the area defines a horizontal plane, and each first electrode of the first electrodes includes a rounded surface oriented toward the horizontal plane.